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J Dent Res 84(6):542-547, 2005
© 2005 International and American Associations for Dental Research

RESEARCH REPORT
Biological

Rotary Culture Enhances Pre-osteoblast Aggregation and Mineralization

S.R. Facer1, R.S. Zaharias2, M.E. Andracki3, J. Lafoon2, S.K. Hunter3, and G.B. Schneider2,4,*

1 Department of Endodontics, University of Iowa, College of Dentistry;
2 Dows Institute of Dental Research, University of Iowa, College of Dentistry, Iowa City, IA 52242, USA;
3 Department of Maternal Fetal Medicine, University of Iowa Health Care and Clinics; and
4 Department of Prosthodontics, University of Iowa, College of Dentistry;

* corresponding author, Galen-Schneider{at}uiowa.edu


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS & METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Three-dimensional environments have been shown to enhance cell aggregation and osteoblast differentiation. Thus, we hypothesized that three-dimensional (3D) growth environments would enhance the mineralization rate of human embryonic palatal mesenchymal (HEPM) pre-osteoblasts. The objective of this study was to investigate the potential use of rotary cell culture systems (RCCS) as a means to enhance the osteogenic potential of pre-osteoblast cells. HEPM cells were cultured in a RCCS to create 3D enviroments. Tissue culture plastic (2D) cultures served as our control. 3D environments promoted three-dimensional aggregate formations. Increased calcium and phosphorus deposition was significantly enhanced three- to 18-fold (P < 0.001) in 3D cultures as compared with 2D environments. 3D cultures mineralized in 1 wk as compared with the 2D cultures, which took 4 wks, a decrease in time of nearly 75%. In conclusion, our studies demonstrated that 3D environments enhanced osteoblast cell aggregation and mineralization.

KEY WORDS: osteoblasts • microgravity • tissue engineering • mineralization


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS & METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Researchers have been investigating the effects of simulated microgravity on cells as a means to study cellular and molecular mechanisms associated with three-dimensional tissue growth (Unsworth and Lelkes, 1998; Botchwey et al., 2001; Rucci et al., 2002; Klement et al., 2004). Three-dimensional growth environment models may provide insight into more physiologically relevant data than two-dimensional systems. Rotary cell culture systems (RCCS) were developed for the analysis of tissue growth in conditions similar to microgravity. It has been shown that the use of RCC allows for the analysis of formation of tissue-like structures in a controlled three-dimensional model. This is due to the alteration of the cell’s perception of a continuously changing gravitational direction as a result of rotational culture conditions (Klement et al., 2004).

When cells are maintained in a 3D growth environment, they tend to aggregate (Granet et al., 1998; Qiu et al., 1998). These types of 3D environments appear to promote cell-cell association while avoiding high shear stress acting on cells (Freed et al., 1999). The horizontal circular rotation allows cells to co-localize and establish a fluid orbit, where there is a high mass transfer of oxygen and nutrients (Botchwey et al., 2001; Rucci et al., 2002). Consequently, cells are suspended in continuous ‘free fall’ at a terminal velocity, with low shear stress force and turbulence (Duray et al., 1997; Freed et al., 1997; Klement et al., 2004). Culturing cells in RCCSs provides an excellent in vitro system for evaluating intercellular and extracellular communications for cell differentiation in a 3D environment similar to that of tissue (Goodwin et al., 1993; Klement et al., 2004).

Previous studies have analyzed the effects of 3D growth environments on osteoblast differentiation using established osteoblast cell lines (Botchwey et al., 2001; Rucci et al., 2002), and have demonstrated increased cellular aggregation and osteoblast differentiation. The conclusions were that the 3D environment may activate compensatory effects on the osteoblasts as a result of negative loading. Those observations led us to hypothesize that 3D environments would enhance the mineralization rate of human embryonic palatal mesenchymal (HEPM) pre-osteoblasts.


   MATERIALS & METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS & METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell Culture
Human embryonic palatal mesenchymal (HEPM 1486; ATCC, Manassas, VA, USA) pre-osteoblast cells were cultured as previously described (Schneider et al., 2004). HEPM pre-osteoblasts (50,000 cells) were plated in micromass cultures, in triplicate, in 10-µL droplets onto plastic tissue culture dishes and placed in a 37°C incubator. The micromass allows the cells in a 2D culture to be maintained in mass rather than dispersed across the tissue culture dish. It allows for a more accurate comparison with those cells cultured in a 3D environment. After 1 hr of attachment, freshly prepared EMEM media treated with ß-glycerophosphate (5 mM) and ascorbate (50 µg/mL) or untreated (no additive) were added to the cultures.

HEPM pre-osteoblasts (10 x 106 cells) were cultured in suspension in a Synthecon RCCS D-410 rotary 10-mL cell culture vessel system (Synthecon, Inc., Houston, TX, USA) and maintained in either the treated or untreated media at an estimated 10–2 g gravity force and 0.5 dynes/cm3 stress force (Unsworth and Lelkes, 1998; Synthecon, Inc., personal communication). All air bubbles were removed from the culture chamber. Cells were initially rotated at 15 rpm. The rotational speed was then increased after aggregates formed and increased in size so that the aggregates would be maintained in suspension. Fresh treated or untreated medium was added biweekly, and cultures were maintained for 1 wk. The media were then removed, and the cultures were gently rinsed with phosphate-buffered saline (PBS) and fixed for 10 min in 10% formalin. Equations of motion governing rotating bioreactors have been previously solved and reported with respect to rotational forces generated within a rotary wall vessel (Pollack et al., 2000; Botchwey et al., 2004).

Hematoxylin & Eosin and Alizarin Red S Staining
HEPM cells were plated in micromass cultures processed at 4 wks. The treated and untreated aggregates were grown in the rotary cell culture, collected at 1 wk, and processed according to standard histological protocols as previously described (Schneider et al., 1999). The 3D specimens were stained with hematoxylin and eosin to depict the specimens’ cellular content and phenotype. Both the 2D and 3D specimens were stained with Alizarin Red S stain for 10 min at room temperature. Wells were washed with nanopure water and evaluated for Alizarin Red S staining as previously described (Schneider et al., 2003), so that we could analyze the specimen’s calcium deposition and location.

Scanning Electron Microscopy (SEM) and Elemental Analysis
We prepared monolayers from the tissue culture plastic cultures and aggregates from the rotary cell cultures for scanning electron microscopy by fixing them in glutaraldehyde in a phosphate buffer. After osmification in 1% OsO4, the samples were dehydrated in ascending concentrations of ethanol for 24 hrs, followed by drying in a critical-point dryer. The samples were then mounted on stubs with copper tape and sputter-coated with carbon rather than gold, to allow for peak analysis of different elements, including calcium and phosphorus, as previously described (Kuempel et al., 1995). Elements were normalized to 100% weight of the sample. Elemental analysis was performed for calcium and phosphorus by energy-dispersive spectroscopy with an EDAX 990 Elemental Detector (EDAX Inc., Mahwah, NJ, USA) in combination with SEM. Triplicate samples were analyzed and weighted for calcium or phosphorus as a percent of total peaks.

Statistical Analysis
Statistical analysis (N = 3) was performed by a one-way analysis of variance (ANOVA) with a Tukey’s Multiple Comparison Test to a confidence level of P < 0.05.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS & METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
2D Pre-osteoblast Cultures Mineralize in 4 Wks
When HEPM pre-osteoblasts were cultured by conventional two-dimensional methods on tissue culture plastic in the absence (Fig. 1AGo) or presence (Fig. 1BGo) of ß-glycerophosphate (ß-GP) and ascorbic acid (AA), no mineralized phenotype was noted at 1 wk. However, when analyzed at 4 wks, the non-treated cultures showed slight mineral deposition, as seen by Alizarin Red S stain for calcium (Fig. 1CGo), and significant mineral deposition when treated with ß-GP and AA (Fig. 1DGo).



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  		Figure 1. Pre-osteoblast monolayer cell cultures on tissue culture plastic mineralized in 4 wks. Alizarin Red S staining for calcium was not evident in cells cultured in the absence (A) or presence (B) of ß-glycerophosphate (ß-GP) and ascorbate at 1 wk. At 4 wks, calcium deposition was greatly enhanced in the presence of ß-GP and ascorbate (D), as compared with non-treated cultures (C). N = 3.

 
Osteoblasts Aggregate in 3D Environments
Although morphologically different, the HEPM pre-osteoblasts formed cellular aggregates in both treated and untreated 3D environments by 1 wk (Fig. 2Go). Use of the RCCS in our studies initiated aggregate formation without beads by the end of the second day in both the treated and the untreated groups of pre-osteoblasts. These small, sand-like masses seen on the second day coalesced to form a larger aggregate by 7 days (Fig. 2Go). However, the treated group aggregate increased in size at a faster rate compared with the untreated group; thus, there was an increase in settling, mandating a faster rotational speed as time progressed. During the biweekly media exchange, requiring the arrestment of rotation, another difference was observed. The treated aggregates fell through the media much like a rock falling through water, as compared with the untreated aggregate, which fell in a gentle, swaying motion, suggesting that there was a difference in density between the two groups. By the end of the week, the aggregates in the treated group outgrew the vessel’s harvesting portal, so we terminated the culture and began our cellular analysis. Gross analysis (Figs. 2AGo, 2BGo) and scanning electron microscopic evaluation (Figs. 2CGo, 2DGo) of the untreated groups revealed a small, compact, cylindrical, and uniform mass (Figs. 2AGo, 2CGo). The treated aggregate (Figs. 2BGo, 2DGo) was larger, and appeared to have microclusters within the larger mass (Fig. 2BGo).



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  		Figure 2. At 1 wk, osteoblast cells cultured in 3D environments had aggregated. Gross and scanning electron microscopy revealed a mass of cells in untreated cultures (A,C) and microclusters within the aggregate in treated cultures (B,D). N = 3. Magnification, 300x. Bar = 50 µm.

 
3D Environments Enhance Osteoblast Mineralization
To determine if the cellular distribution within the aggregates of both the treated and untreated cultures differed at 1 wk, we performed histological analysis using hematoxylin and eosin staining. This assay revealed that cells were distributed across both the untreated (Fig. 3AGo) and treated (Fig. 3BGo) aggregates. However, within the aggregates, the patterns of cell distribution differed. In the untreated aggregate, the mass was uniform (Fig. 2CGo), whereas the treated aggregate was made up of smaller aggregates (Fig. 2DGo). The patterns of cell distribution were similar. In the untreated 3D culture (Fig. 3AGo), cells were uniformly distributed across the aggregate. In the treated 3D cultures, the cells were distributed within the smaller aggregates (Fig. 3BGo) that made up the larger aggregate. The cellular pattern within the treated aggregate (Fig. 3BGo) mimicked the microclustered morphology seen in Fig. 2DGo.



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  		Figure 3. Enhanced mineralization was observed in cultures maintained in 3D environments for 1 wk. Uniform cell distribution and minimal calcium and phosphorus deposition were noted in non-treated cultures (A,C,E). When treated with ß-GP and ascorbate, the cell distribution and enhanced mineralization pattern matched the pattern of microclusters within the aggregate (B,D,E). Magnification, 20x. Bar = 100 µm. Asterisks (*) indicate significant differences between conditions (N = 3) at P < 0.05.

 
Differences in patterns of cellular distribution within the aggregates led us to question whether the 3D environment might also affect the pattern of mineralization as well. Differences in patterns of mineralization were noted between pre-osteoblasts cultured in 3D environments at 1 wk as compared with those cultured in 2D conditions (Fig. 3Go). In addition, Alizarin Red S staining showed that calcium deposition was increased at 1 wk in treated cultures maintained in 3D environments (Fig. 3DGo), as compared with non-treated aggregates (Fig. 3CGo). The calcium deposition in the treated aggregates (Fig. 3DGo) was localized to the microclusters in a pattern similar to that seen with the cellular distribution in Fig. 3BGo. The levels of calcium noted by Alizarin Red S stain in the non-treated 3D environment aggregates were minimal.

Quantitative levels of calcium and phosphorus were detected by energy-dispersive spectroscopic elemental analysis in combination with scanning electron microscopy (Fig. 3EGo). No significant differences were noted in calcium or phosphorus levels in cultures maintained for 1 wk in 2D cultures. However, at 1 wk, significant (P < 0.001) three- to 18-fold increases in levels of both calcium and phosphorus were detected in the 3D cultures. Levels of calcium in the treated 3D cultures were three-fold higher (P < 0.001) when compared with those in untreated 3D cultures. Analysis of cell number and protein concentration at 1 wk of both 2D and 3D cultures revealed that, by 1 wk, the 3D cultures had an 85% decrease in cell number, whereas the 2D cultures had a nearly 95% increase in cell number, yet the 3D cultures still had an 18-fold higher level of calcium. In addition, mineral was detected in the treated 3D cultures at 1 wk, as compared with the treated 2D cultures, in which mineral was not detected until 4 wks. This suggests that osteoblasts cultured in a 3D environment will mineralize nearly 75% faster than those that are cultured in a 2D environment but otherwise treated the same.

3D Aggregate Cells Grow in 2D Culture Conditions
Knowing that the cell number decreased in the 3D environment aggregates, we wanted to determine if the cells remaining in the aggregate still had the potential to grow if re-introduced into a conventional 2D tissue culture environment (Fig. 4Go). When compared with normal 2D tissue-cultured cells, or cells isolated from 3D environments prior to aggregation (Fig. 4AGo), the 3D aggregates (Fig. 4BGo) demonstrated normal cellular growth within 24 hrs of explantation in the 2D environment (Fig. 4BGo, arrow). Cells continued to grow until reaching confluence and then, through normal cellular processes, became senescent as a result of contact inhibition (Folkman and Moscona, 1978), suggesting that the aggregate cells had not transformed.



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  		Figure 4. 3D pre-aggregated cells (A) or aggregates (B) when re-introduced into 2D tissue culture conditions had cellular outgrowth (B, arrow) mimicking that of normal tissue cultured cells. N = 3. Magnification, 10x. Bar = 10 µm.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS & METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The horizontal rotation of the rotary cell culture inhibits cells from adhering within the vessel, although cells do adhere to the extracellular matrix created within the aggregate (Klement et al., 2004). This system enhances cell aggregation by maintaining the cells in a fluid orbit, which in effect keeps the cells in a constant state of free fall. This environment creates an opportunity for tissue development to be studied in three-dimensional environments. As has been reported, the cells’ perception of downward gravity is altered during rotation, since the gravitational vectors are continuously re-oriented as a function of rotation (Klaus, 2001).

Mineralization of the pre-osteoblasts was significantly increased in only the treated 3D cultures, suggesting that there was an enhanced effect on differentiation to a mineralized phenotype. This occurred by 7 days, as compared with the tissue plastic cultures grown according to standard methods, which took nearly 4 wks to mineralize. The aggregates from the non-treated microgravity cultures showed slight evidence of increased calcium and phosphorus at 1 wk, but not nearly to the level of that seen in the treated 3D conditions. These findings were very interesting in light of the fact that, typically, when normal cell-to-extracellular-matrix interactions are disrupted, the cells die.

Typically, when cells become suspended, they undergo anoikis, apoptosis as a result of lost cell adhesion to the underlying matrix (Attwell et al., 2000). The association of multi-potent mesenchymal cells with the extracellular matrix mediates the differentiation of the cells into osteoblasts (Weissman et al., 2001). We have shown that adhesion-mediated events through integrin receptors, in particular {alpha}2ß1, and its interaction with the extracellular matrix are important and necessary for the mediation of osteoblast differentiation (Schneider et al., 2001). Previous studies have attempted to analyze molecular mechanisms associated with osteoblast differentiation by the use of three-dimensional osteoblast cell model systems cultured onto tissue culture plastic (Cooper, 1998; Schneider et al., 1999). However, few studies have analyzed the effects of 3D environments on osteoblast cell behavior. Some reports have described the aggregation of cells when cultured in 3D environments alone (Rucci et al., 2002) or on scaffolds (Botchwey et al., 2001). These studies describe the enhanced rate and ability of the cells to mineralize, and demonstrated enhanced expression of osteoblast differentiation markers such as alkaline phosphatase, bone sialoprotein, and osteonectin. We found similar results with the HEPM pre-osteoblasts when cultured in 3D environments. The HEPM cells began to aggregate within 2 days, and by day 7, the cultures, when treated with ß-glycerophosphate and ascorbate, mineralized. Mineralization of 3D cultures was detected at 1 wk, whereas the 2D cultures took 4 wks to initiate mineralization, even in the presence of ß-glycerophosphate and ascorbate, suggesting that the conditions of the 3D environment enhanced the molecular mechanisms associated with mineralization initiation as cell numbers and possible density decreased.

The precise cellular and molecular mechanisms for adhesion-mediated osteoblast differentiation in 3D environments remain undefined. Our studies, as well as those of others (Botchwey et al., 2001; Rucci et al., 2002; Klement et al., 2004), suggest that the 3D environment may enhance the mechanisms of osteoblast cell biology. The environment may activate alternative pathways that are more closely related to physiologic conditions. We have demonstrated that, in a three-dimensional fetal bovine mandibular osteoblast model, the spatiotemporal expressions of key integrins and associated extracellular matrix proteins were similar in timing and location of expression (Schneider et al., 1999). This same type of phenomenon may occur in simulated 3D environments. As others (Rucci et al., 2002) have discussed, enhanced mineralization may be associated with a compensatory effect by the osteoblast at the cellular level as a result of 3D growth. Or it may be that when cells aggregate, they interact with their surrounding extracellular matrix via integrin receptors and thus are maintained in an environment similar to that found in vivo in normal tissues. This would allow the cells to interpret environmental cues from the matrix and initiate osteogenesis without sensing the loss of adhesion as a result of altered gravity vectors.

The exact cellular and molecular mechanisms affecting osteoblast differentiation under simulated microgravity remain poorly defined. Our findings demonstrate that HEPM pre-osteoblasts mineralize rapidly as compared with current conventional tissue culture methods. The behavior of the cells and aggregates grown in the RCC demonstrates what appears to be normal, but accelerated, growth and differentiation. This may then translate into the development of novel tissue-engineering strategies, where rotary cell culturing may be used not only as a tool for study, but also as a tool for the development of osseous tissue-engineering strategies for the grafting of craniofacial defects and deficiencies.


   ACKNOWLEDGMENTS
 
This work was supported by grants R03DE014269 (GS) and P60DE13076 (GS).

Received April 19, 2004; Last revision February 22, 2005; Accepted March 22, 2005


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS & METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Attwell S, Roskelley C, Dedhar S (2000). The integrin-linked kinase (ILK) suppresses anoikis. Oncogene 19:3811–3815.[ISI][Medline]

Botchwey EA, Pollack SR, Levine EM, Laurencin CT (2001). Bone tissue engineering in a rotating bioreactor using a microcarrier matrix system. J Biomed Mater Res 55:242–253.[ISI][Medline]

Botchwey EA, Pollack SR, Levine EM, Johnston ED, Laurencin CT (2004). Quantitative analysis of three-dimensional fluid flow in rotating bioreactors for tissue engineering. J Biomed Mater Res 69(A):205–215.

Cooper LF (1998). Biologic determinants of bone formation for osseointegration: clues for future clinical improvements. J Prosthet Dent 80:439–449.[ISI][Medline]

Duray PH, Hatfill SJ, Pellis NR (1997). Tissue culture in microgravity. Sci Med (Phila) 4(3):46–55.

Folkman J, Moscona A (1978). Role of cell shape in growth control. Nature 273:345–349.[Medline]

Freed LE, Langer R, Martin I, Pellis NR, Vunjak-Novakovic G (1997). Tissue engineering of cartilage in space. Proc Natl Acad Sci USA 94:13885–13890.[Abstract/Free Full Text]

Freed LE, Pellis N, Searby N, de Luis J, Preda C, Bordonaro J, et al. (1999). Microgravity cultivation of cells and tissues. Gravit Space Biol Bull 12(2):57–66.[Medline]

Goodwin TJ, Schroeder WF, Wolf DA, Moyer MP (1993). Rotating-wall vessel coculture of small intestine as a prelude to tissue modeling: aspects of simulated microgravity. Proc Soc Exp Biol Med 202:181–192.[Abstract]

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